Immunoassay for Detection of Virus-Antibody Nanocomplexes in Solution by Chip-Based Pillar Array

ABSTRACT

Techniques for detection of virus-antibody nanocomplexes using a chip-based pillar array are provided. In one aspect, a method for virus detection is provided. The method includes the steps of: collecting a fluid sample from a virus-bearing source; contacting the fluid sample with an antibody that binds to viruses to form a sample-antibody mixture, wherein the antibody is labeled with a fluorescent tag; separating particles including any antibody-virus complexes, if present, from the sample-antibody mixture using an assay nanopillar array; and detecting the antibody-virus complexes, if present, in the particles from the separating step using fluorescence. A virus detection chip device and a chip-based immunoassay method are also provided.

FIELD OF THE INVENTION

The present invention relates to virus detections, and moreparticularly, techniques for detection of virus-antibody nanocomplexesusing a chip-based pillar array.

BACKGROUND OF THE INVENTION

Viruses replicate and spread through infection of a host organism. Fordisease-causing viruses, controlling the spread of a viral infection iscause for global concern. Early detection of a viral infection is key toeffectively treating infected patients and preventing epidemic spread.

The incubation period for many viruses is on the order of days to weeksand the infection period often begins before symptoms are present. It istherefore often challenging to prevent patient-to-patient infectionwithout sensitive methods to detect viral infection before symptomsappear. Additionally, the effectiveness of many of the availableantiviral medications rely on early detection and treatment of disease.

Although technology exists for detection of viral infection, thesensitivity is limited and most rely on sophisticated laboratoryequipment and trained technicians. Thus in order to reduce the number ofviral infections and improve early treatment, technology is needed forrapid point-of-care viral infection detection by individuals with littlemedical training.

Current methods for the detection of viral infections in patients relyprimarily on “wet” lab techniques, which require an appropriatelyequipped laboratory and trained staff. Turnaround time for even the mostrapid tests is often hours or days, limiting their use as point-of-careviral detection methods. Two commonly used techniques are enzyme-linkedimmunosorbent assay (ELISA) and polymerase chain reaction (PCR).

ELISA tests detect virus in biological samples through the use ofantibodies that bind viral antigens. Viral particles are adhered to asolid surface such as a plastic 96-well plate and are subsequently boundby antibodies that recognize a viral antigen. These primary antibodiesor secondary antibodies are covalently linked to a fluorescent moleculeor enzyme that produces colorimetric or luminescent signal upon exposureto a ligand. This signal is measured by a fluorimeter orspectrophotometer. ELISA can also be adapted to detect antibodiesdeveloped against virus in patient samples, however antibodies againstvirus are produced later in the infection process.

Although some ELISA methods are considered highly sensitive and can bequantitative, they require expensive laboratory equipment and theirreproducibility is dependent on a well-trained technician. They are notsuitable for rapid point-of-care detection and often require severalhundred microliters of sample. Additionally, there is often disagreementover the numerical cutoff value of the quantified signal leading topotential false positives and false negatives.

PCR and reverse transcription PCR (RT-PCR) detect the nucleic acidcomponent, i.e., deoxyribonucleic acid (DNA) or ribonucleic acid (RNA),of the virus and are generally considered to be more sensitive thanELISA with fewer false positives. PCR uses RNA primers designed to matchknown sequences in the viral RNA or DNA to amplify fragments which areused as a readout for presence of the virus. PCR and RT-PCR can bequantitative, allowing for accurate calculation of viral load. However,both PCR-based methods require sophisticated and expensive laboratoryequipment and trained technicians, eliminating their use as rapidpoint-of-care tests.

A handful of rapid point-of-care diagnostic tests exist, with the twomost widely used being for the detection of human immunodeficiency virus(HIV) and influenza A and B. The advantage of these tests is their easeof use and rapid results, usually in less than 30 minutes. However, theHIV rapid test relies on the detection of antibodies generated by thepatient against HIV, which can take several months to reach a detectablelevel. In line with this, these tests are only qualitative in nature andhave been shown to have a high occurrence of false positive and falsenegative results.

Therefore, there is a need for sensitive, quantitative detection ofviral particles in a rapid point-of-care protocol amenable toself-administration with small sample volumes.

SUMMARY OF THE INVENTION

The present invention provides techniques for detection ofvirus-antibody nanocomplexes using a chip-based pillar array. In oneaspect of the invention, a method for virus detection is provided. Themethod includes the steps of: collecting a fluid sample from avirus-bearing source; contacting the fluid sample with an antibody thatbinds to viruses to form a sample-antibody mixture, wherein the antibodyis labeled with a fluorescent tag; separating particles including anyantibody-virus complexes, if present, from the sample-antibody mixtureusing an assay nanopillar array; and detecting the antibody-viruscomplexes, if present, in the particles from the separating step usingfluorescence.

In another aspect of the invention, a virus detection chip device isprovided. The virus detection chip device includes: a capillary openingfor accepting a fluid sample collected from a virus-bearing source; amixing reservoir, connected to the capillary opening, for contacting thefluid sample with an antibody that binds to viruses, wherein theantibody is labeled with a fluorescent tag to form a sample-antibodymixture in cases where the antibody and sample are not pre-mixed; afirst filtering nanopillar array, connected to the mixing reservoir, forremoving particles from the sample-antibody mixture; a second nanopillararray, connected to the first nanopillar array, for separating particlesincluding any antibody-virus complexes, if present, from thesample-antibody mixture; and a diode-induced fluorescence detector,connected to the second nanopillar array, for detecting theantibody-virus complexes, if present, in the particles usingfluorescence.

In yet another aspect of the invention, a chip-based immunoassay methodis provided. The method includes the steps of: providing a virusdetection chip device having: a capillary opening; a mixing reservoirconnected to the capillary opening; a first nanopillar array connectedto the mixing reservoir; a second nanopillar array connected to thefirst nanopillar array; and a diode-induced fluorescence detectorconnected to the second nanopillar array; introducing a fluid samplecollected from a virus-bearing source to the virus detection chip devicethrough the capillary opening; in the mixing reservoir, contacting thefluid sample with an antibody that binds to viruses, wherein theantibody is labeled with a fluorescent tag to form a sample-antibodymixture; removing particles from the sample-antibody mixture using thefirst nanopillar array; separating particles including anyantibody-virus complexes, if present, from the sample-antibody mixtureusing the second nanopillar array; and detecting the antibody-viruscomplexes, if present, in the particles using the diode-inducedfluorescence detector.

A more complete understanding of the present invention, as well asfurther features and advantages of the present invention, will beobtained by reference to the following detailed description anddrawings.

BRIEF DESCRIPTION OF THE DRAWINGS

FIG. 1 is a diagram illustrating an exemplary methodology for virusdetection according to an embodiment of the present invention;

FIG. 2 is a schematic diagram illustrating use of the present virusdetection chip device in performing the methodology of FIG. 1 accordingto an embodiment of the present invention;

FIG. 3a is a schematic diagram of a nanopillar array illustrating thearray parameters θ_(max), G, and pillar pitch, λ according to anembodiment of the present invention;

FIG. 3b is a scanning electron microscope (SEM) image of a sorting arrayaccording to an embodiment of the present invention;

FIG. 3c is a table showing the nanopillar array gap sizes G and measuredpercentage maximum angle P in nanopillar arrays, for a given particlediameter D_(P), at flow velocities between 200-350 μm/s according to anembodiment of the present invention;

FIG. 3d is an image illustrating lateral displacement modes for zigzag,partial, and bumping according to an embodiment of the presentinvention;

FIG. 3e are SEM images of inlet and outlet regions bordering the arrayaccording to an embodiment of the present invention;

FIG. 3f are fluorescence microscopy images of fluorescent polystyrenebeads flowing into the inlet region (top row) and exiting the arrayoutlet region (bottom row), corresponding to those shown in the SEMimages in FIG. 3e . according to an embodiment of the present invention;

FIG. 4a is a schematic diagram of an H1N1 viral particle withfluorescence detection scheme according to an embodiment of the presentinvention;

FIG. 4b is a composite fluorescence microscopy image showingdisplacement of H1N1 viral particles in a nanoscale deterministiclateral displacement array according to an embodiment of the presentinvention;

FIG. 4c is a histogram of percentage maximum angle across the outlet ofthe array shown in FIG. 4b according to an embodiment of the presentinvention;

FIG. 4d is a schematic diagram of a human placental exosome withfluorescence detection scheme according to an embodiment of the presentinvention;

FIG. 4e is a composite fluorescence microscopy image showingdisplacement of exosomes in a nanoscale deterministic lateraldisplacement array according to an embodiment of the present invention;

FIG. 4f is a histogram of percentage maximum angle across the outlet ofthe array shown in FIG. 4e according to an embodiment of the presentinvention;

FIG. 5a are fluorescence image mosaics of double stranded DNA (dsDNA)displacement in a deterministic lateral displacement array according toan embodiment of the present invention;

FIG. 5b are normalized fluorescence intensity line profiles of displacedDNA taken at the outlet of each array according to an embodiment of thepresent invention;

FIG. 5c is a diagram illustrating the percentage maximum angle of dsDNAas a function of DNA length according to an embodiment of the presentinvention;

FIG. 6a is an image of fluidic chips printed on a wafer according to anembodiment of the present invention;

FIG. 6b is an optical image showing microfluidic channels joined bynanochannel features, including pillar sorting arrays, according to anembodiment of the present invention;

FIG. 6c is a Fast Fourier Transform (FFT) confirming successfulpatterning of the design angle θ_(t) according to an embodiment of thepresent invention;

FIG. 6d is an SEM image of a sorting array according to an embodiment ofthe present invention;

FIG. 7a is an SEM image of D_(P)=110 nm beads according to an embodimentof the present invention;

FIG. 7b is an SEM image of D_(P)=50 nm beads according to an embodimentof the present invention;

FIG. 7c is an SEM image of D_(P)=20 nm beads according to an embodimentof the present invention;

FIG. 7d is a histogram of bead diameters measured from the SEM images inFIGS. 7a-c according to an embodiment of the present invention;

FIG. 7e is a table of properties of the bead samples according to anembodiment of the present invention;

FIG. 8a is a schematic diagram of an exemplary custom fluid jigaccording to an embodiment of the present invention;

FIG. 8b is a top-down image of the custom fluid jig with a nanoscaledeterministic lateral displacement chip loaded according to anembodiment of the present invention;

FIG. 9a is a schematic diagram of a nanoscale deterministic lateraldiffusion array showing particle flux entering from left (inlet) toright (outlet) according to an embodiment of the present invention;

FIG. 9b is a diagram of a plot of fluorescent line profiles taken at theoutlet of arrays for a complete displacement scenario according to anembodiment of the present invention;

FIG. 9c is a diagram of a plot of fluorescent line profiles taken at theoutlet of arrays for a partial displacement scenario according to anembodiment of the present invention;

FIG. 10 is a diagram illustrating polystyrene fluorescent beaddisplacement as a function of particle diameter compared to criticaldiameter needed for displacement in a parabolic flow according to anembodiment of the present invention;

FIG. 11 is a diagram illustrating percentage maximum angle offluorescent polystyrene beads displaced in nanoscale deterministiclateral displacement arrays as a function of the scaling ratio betweenparticle diameter and gap size according to an embodiment of the presentinvention;

FIG. 12 is a table of performance parameters for nanoscale deterministiclateral displacement of fluorescence polystyrene beads according to anembodiment of the present invention;

FIG. 13a is a fluorescent microscopy image of the trajectory of anexosome through the nanoscale deterministic lateral displacement arrayparticle according to an embodiment of the present invention;

FIG. 13b is a diagram illustrating collection of single-particle exosometrajectories taken at the outlet of a G=235 nm array according to anembodiment of the present invention;

FIG. 13c is a histogram of single-particle positions at the outletaccording to an embodiment of the present invention;

FIG. 14a is an image of viral particle experimental controls whichillustrates that virus alone shows no fluorescence according to anembodiment of the present invention;

FIG. 14b is an image of viral particle experimental controls whichillustrates that antibodies alone follow the streamlines of the laminarflow according to an embodiment of the present invention;

FIG. 14c is an image of viral particle experimental controls whichillustrates that anti-H1N1 antibody-virus complexes are bumped accordingto an embodiment of the present invention;

FIG. 14d is an image of viral particle experimental controls whichillustrates that non-specific M13 antibodies do not bind viral particlesand therefore do not bump according to an embodiment of the presentinvention; and

FIG. 15 is a table of end-to-end coil diameters and scaling ratioscalculated for dsDNA in nanoscale deterministic lateral displacementexperiments according to an embodiment of the present invention.

DETAILED DESCRIPTION OF PREFERRED EMBODIMENTS

Provided herein are nanotechnology-based techniques for sensitive,quantitative detection of viral particles from biological samples (e.g.,blood, saliva, sweat, urine, plant samples, food samples, and drinkingwater) using antibodies against viral antigens in a rapid point-of-careprotocol. Advantageously, the present techniques are carried out via aone-step system (thus simplifying the detection process), which could beself-administered and require only a small biological sample, similar toglucose monitoring systems used by diabetics.

Namely, as will be described in detail below, the present techniquesutilize a silicon (Si) nanotechnology-based approach to identifyfluorescently-labeled antibodies bound to viral particles in abiological sample, allowing for the detection of virus in a patient. Asmall-volume of sample (blood, saliva, sweat, etc.) would be incubatedwith a fluorescently-labeled antibody against the suspected virus. Themixed sample is applied to a diagnostic silicon chip (lab-on-a-chip(LOC)) device which uses an array of nanopillars to sort biomoleculesbased on size. Unbound antibodies are too small to be sorted and flowthrough the chip, while antibodies bound to virus are above the sizethreshold and are sorted. These sorted particles can be detected andquantified directly by fluorescence microscopy, e.g., by a fluorimeteror by an on-chip diode-induced fluorescence detector.

The primary advantages of the present virus detection chip over currentviral detection methods include: smaller sample volume, lack ofrequirement for sophisticated laboratory equipment, portability,self-administration by an untrained individual, adaptability to anyvirus with an available antibody, and early detection capabilities. Asdescribed above, current viral detection technologies includeenzyme-linked immunosorbent assay (ELISA) and polymerase chain reaction(PCR). Both of these techniques require well-equipped laboratories andtrained technicians and can take hours to days to produce results,making them unsuitable for point-of-care diagnosis.

The present virus detection chip device is compact and compatible withon-chip detection technology making it highly portable. It requires lessthan 100 uL of sample volume and could be implemented in a mannersimilar to glucose monitoring kits for diabetics, makingself-administration by a patient possible. The present virus detectionrelies on a fluorescent signal from an antibody to a specific virus. Asantibodies exist for most viral capsids or envelopes, it is possible todesign a chip to test for the presence of virtually any virus. Furtherthe detection protocol described herein is not limited to human samples,but could also be used to detect plant and animal viruses providedantibodies are available.

Finally, many conventional virus detection tests rely on the productionof viral antibodies in the patient, which occurs later in diseaseprogression. The present techniques are sensitive enough to detect asmall number of viral particles directly and could be used to determinean infection early in progression.

The present techniques are now described in detail by way of referenceto methodology 100 of FIG. 1. In step 102, a fluid sample is collected.In general, fluid samples can be collected from virtually anyvirus-bearing source. This includes, but is not limited to, blood,saliva, sweat, plant tissue, drinking water, and food products.According to an exemplary embodiment, less than 100 microliters (μl)(e.g., from about 50 μl to about 100 μl, and ranges therebetween) is allthat is needed to be collected for testing, since only a small amount ofthe fluid sample needs to be introduced to the capillary opening of thepresent (disposable) virus detection chip device (see below).

Non-blood samples could be collected at the point-of-care or in thefield for rapid testing. Blood samples could also be produced quicklyusing a disposable lancet at the point-of care. This test could be usedfor any virus in the 100 nanometer (nm) size range, or larger, that hasantibodies available for binding to the outer capsid or envelope of thevirus. This includes, but is not limited to influenza viruses,adenoviruses, Ebola and Marburg viruses, poxviruses (including smallpox), and herpes viruses (including Epstein-Barr virus andVaricella-zoster virus).

The fluid sample collected in step 102, is then prepared for analysis.It is notable however that only minimal sample preparation is needed,and most of the preparative steps can (if so desired) be built into thevirus detection chip device itself. In general, to prepare the samplefor analysis, the sample is contacted (i.e., mixed) with afluorescently-labeled antibody that will bind to the virus capsid orenvelope.

A variety of fluorescent tags could be used and include, but are notlimited to, quantum dots, Alexa Fluors® (available from LifeTechnologies™, Grand Island, N.Y.), fluorescein, rhodamine, Oregongreen, pyrene, and HyLite™ Fluor dyes (available from AnaSpec, Inc.,Fremont, Calif.). Antibodies will be conjugated to these fluorescenttags using covalent linkages including, but not limited to, amino,carboxyl, thiol, and azide chemistries.

According to an exemplary embodiment, the sample preparation is carriedout by first mixing the fluid sample with a solution containing thefluorescently-labeled antibody. See step 104. The sample/antibodymixture can then be introduced to the capillary opening of the virusdetection chip device in step 106.

The present virus detection chip device, however, can have a built-inloading reservoir (see below) in which the tagged antibody can belyophilized/dried on the chip so that there is no mixing required forthe sample to be loaded on the chip—i.e., the sample simply gets loadedand mixes on chip with the antibody. This alternative embodiment wherethe sample preparation steps are built into the chip itself simplifiesthe assay process, which can be beneficial for point-of-care access tothe present techniques.

By way of example only, to pre-load the chip with tagged antibody atleast one picogram of antibody is lyophilized in the loading reservoirof the chip. See step 108. This amount of antibody is in excess of thevirus and will ensure that each viral particle can be bound by severalantibodies assuming a viral load of over 10,000 copies/milliliter. Ofcourse, lower viral loads can also be detected, but will simply be boundmy more antibodies until each virus is saturated by bound antibody.

In step 110, the sample (collected in step 102) is introduced to thecapillary opening of the present virus detection chip device. A samplebuffer of phosphate-buffered saline (PBS, 10 millimolar (mM) sodiumphosphate, 18 mM potassium phosphate, 137 mM sodium chloride, 2.7 mMpotassium phosphate) is preferably loaded behind the sample to ensurethat the entire sample enters the array. Following flow through thecapillary opening the sample will enter the loading reservoir, where itwill mix and bind with the antibody (step 112) prior to entering thefiltration and pillar arrays. With whichever procedure is implemented,i.e., either premixing the sample with a tagged antibody solution (steps104-106) or passing the sample through the tagged antibody via theloading reservoir of the chip (steps 108-112), a mixture containing thesample and tagged antibody is now present within the virus detectionchip device.

As will be described in detail below, the virus detection chip deviceemployed herein contains an array of nanopillars, through which thesample/antibody mixture will pass, which will serve to separate (bysize) antibody-virus complexes present in the sample/antibody mixture.Thus, this nanopillar array may also be referred to herein as an “assaynanopillar array.”

The sample/antibody mixture might also contain larger particles (e.g.,particles with dimensions greater than 500 nm), such as organelles, cellmembrane, and protein aggregates. These larger particles can clog theassay nanopillar array, and thus it is desirable to remove them from thesample before the sample passes through the assay nanopillar array. Seestep 114. Therefore, the virus detection chip device preferably containsanother array of nanopillars (also referred to herein as a “filteringnanopillar array”) before the assay nanopillar array which serves tofilter out (i.e., remove) these larger particles from the sample.Namely, the sample/antibody mixture passes first through the filteringnanopillar array wherein particles larger than 500 nm are removed. Thesample then passes through the assay nanopillar array whereinantibody-virus complexes, if present, are separated out from themixture. See step 116. The filtering nanopillar array and the assaynanopillar array may also be referred to herein as a “first” and a“second” nanopillar array, respectively. Further, as will be describedin detail below, the filtering nanopillar array and the assay nanopillararray differ generally in the size/spacing of the nanopillars in thearray.

The filtering and assay nanopillar arrays implemented herein are, whatis known in the art as, deterministic lateral displacement pillararrays. Deterministic lateral displacement pillar arrays in silicon haveproven an efficient technology to sort, separate, and enrichmicrometer-scale particles. See, for example, Huang et al., “ContinuousParticle Separation Through Deterministic Lateral Displacement,”Science, vol. 304 (May 2004) (hereinafter “Huang”); Inglis, et al.,“Critical particle size for fractionation by deterministic lateraldisplacement,” Lab Chip, 6, 655-658 (March 2006) (hereinafter “Inglis”);and Loutherback et al., “Improved performance of deterministic lateraldisplacement arrays with triangular posts,” Microfluid Nanofluid9:1143-1149 (May 2010), the contents of each of which are incorporatedby reference as if fully set forth herein. However, deterministiclateral displacement pillar array technology has never been translatedto the nanoscale.

According to an exemplary embodiment, the filtering nanopillar array isa 1 micrometer (μm) gap (between the pillars) nanopillar array whichsorts particles larger than 500 nm in size and “bumps” them to the rightside of the array. The principles behind the mechanism of bumping aredescribed, for example, in Huang and Inglis. The size of particlessorted is dependent on the ratio of the spacing between the nanopillarsand the size of the nanopillars themselves. These larger particles willbe sorted in step 114 by the filtering nanopillar array into a wastereservoir on the right side of the chip (see below).

After removal of large particles, the sample will enter the assaynanopillar array (e.g., a 120 nm nanopillar array) which will separateantibody-virus complexes, if present, from the mixture. See step 116.Namely, all particles in the mixture that are larger than about 100 nmwill be sorted by the assay nanopillar array and bumped to the right,including the virus bound by the fluorescent antibody.

In step 118, fluorescence can then be used to detect whether or notantibody-virus complexes are present in the particles retrieved in step116. Again, this detection functionality can be built directly into thechip. For example, particles that exit the right side of the assaynanopillar array (see above) can be collected in an on-chipdiode-induced fluorescence detector, which can detect and quantifyfluorescent signal from the fluorescently labeled antibody-viruscomplex. Any signal detected above a pre-calibrated threshold willindicate the presence of virus in the sample. Any background particleslarger than 100 nm that enter the 120 nm assay nanopillar array will notaffect detection as they will not be fluorescently labeled and will notproduce a signal.

In a sample lacking the virus detected by the antibody, the antibodyalone will not be deflected by the assay nanopillar array. Antibodiesconsist of two large and two small chains which total 160 kilo-Daltonsin molecular weight. Assuming a globular shape, each unbound antibodywould have an estimated size of less than 10 nm, and would flow straightthrough the 120 nm assay nanopillar array rather than being bumped tothe right. Therefore, no fluorescent signal will be collected ordetected by the diode-induced fluorescence detector as all of thefluorescently-labeled antibodies will flow out the bottom of the chip.

FIG. 2 is a schematic diagram illustrating use of the above-described(disposable) virus detection chip device in performing methodology 100of FIG. 1. For illustrative purposes, a scenario where the sample doesnot contain the virus is shown on the left side of the figure and ascenario where the sample contains the virus is shown on the right sideof the figure. A fluid sample first must be collected. As describedabove, the fluid sample used herein can be collected from anyvirus-bearing source, such as blood, saliva, sweat, plant tissue,drinking water, and food products. Further, only a small amount of thefluid sample is needed for testing, for example, less than 100 μl (e.g.,from about 50 μl to about 100 μl, and ranges therebetween)—see above.The presence of the virus in the right hand sample is indicated using amulti-faceted polygon (i.e., to represent the virus capsid).

As described above, the fluid sample can (optionally) be premixed with afluorescently-tagged antibody solution or, as shown in FIG. 2,lyophilized/dried tagged antibody can be placed in the mixing reservoirof the chip. In the latter case, the sample itself can be collected bythe capillary opening of the chip. The capillary opening is connected tothe mixing reservoir. Thus, from the capillary opening, the sample willflow through the mixing reservoir where the tagged antibody will bind tothe virus, if present, in the sample. The result is a sample-antibodymixture. As shown in FIG. 2, in the case where the virus is not presentin the sample (example shown on the left) the mixing reservoir willcontain no virus-antibody complexes (i.e., all that is shown in themixing reservoir in the “No Virus” scenario is the fluorescent taggedantibody). On the other hand, in the case where the virus is present inthe sample (example shown on the right) the mixing reservoir willcontain virus-antibody complexes (i.e., virus-antibody complexes areshown in the mixing reservoir in the “Virus” scenario).

As provided above, a sample buffer (e.g., PBS) can be loaded behind thesample to ensure that all of the sample enters the nanopillar arrays.Namely, the mixing reservoir is connected to the (first) filteringnanopillar array. As provided above, the filtering array serves toremove larger particles from the sample-antibody mixture that canpotentially clog the smaller assay nanopillar array. As shown in FIG. 2,the particles filtered out of the mixture by the filtering nanopillararray are sent off as waste (labeled “Waste (>500 nm)”) through aconduit off to the right side of the chip.

The mixture then moves to the (second) assay nanopillar array which, asshown in FIG. 2, is connected to the (first) filtering nanopillar array.As provided above, the assay nanopillar array will separateantibody-virus complexes, if present, from the mixture. Namely, allparticles in the solution that are larger than about 100 nm will besorted by the assay nanopillar array and bumped to the right, includingthe virus bound by the fluorescent antibody. Thus, as shown in FIG. 2,in the case where the virus is not present in the sample (example shownon the left) no virus-antibody complexes are present, and as such novirus-antibody complexes are bumped (i.e., separated out) by the assaynanopillar array. It is notable that the mixture might in fact containparticles (other than virus-antibody complexes) that might get separatedout at this stage (and which are not shown in FIG. 2). However, theseparticles will not be fluorescently tagged, and thus will not contributeto the detection carried out at the diode (i.e., as shown in FIG. 2unbound tagged antibodies will pass through the chip as waste). On theother hand, in the case where the virus is present in the sample(example shown on the right) virus-antibody complexes are present andwill be bumped (i.e., separated out) by the assay nanopillar array. Seevirus-antibody complex directed (via the assay nanopillar array) to theconduit leading to the diode-induced fluorescence detector (labeled“Diode”).

Finally, the diode-induced fluorescence detector which, as shown in FIG.2, is used to detect and quantify the presence of virus-antibodycomplexes in the particles separated out by the assay nanopillararray—due to the presence of the fluorescent tags. As provided above,any signal detected above a pre-calibrated threshold can be used toindicate the presence of virus in the sample. Any background particleslarger than 100 nm that enter the 120 nm assay nanopillar array will notaffect detection as they will not be fluorescently labeled and will notproduce a signal.

The present nanopillar array-based techniques can be used to rapidlysort biological entities down to 20 nm in continuous flow, usingnanoliter volumes, with single-particle resolution. Thus, in addition toviruses, the present techniques open the possibility for sorting a widerange of biological entities, setting the foundation for novelapplications in single-cell fractionation, proteomics and point-of-caremedical diagnostics. For example, the ability to sort exosomes, secretedlipid vesicles with sizes between 30 nm to 120 nm containing a proteinand nucleic acid cellular snapshot have been demonstrated in accordancewith the present techniques. The ability to sort exosomes based on sizeand surface markers is important to future medical applications, andsuggests the viability of sorting a variety of lipid membrane basedparticles such as synaptic vesicles. Further, separation of doublestranded DNA (dsDNA) fragments by length at the single molecule levelhas been demonstrated in accordance with the present techniques, whichhas broad applications in genomic sequencing and epigenetics.

One important parameter in the study of deterministic lateraldisplacement pillar arrays is the Peclet number, defined as the ratio ofthe time for a particle to diffuse a characteristic distance d in thesystem to that required to advect the same distance, which decreasesrapidly as the feature size decreases at fixed fluid speed. The Pecletnumber is proportional to d and the fluid speed. The concern forparticle sorting in deterministic lateral displacement is that at toolow a Peclet number, diffusion will overwhelm the displacement processand the motion will no longer be deterministic. Therefore one goalherein is to determine the behavior of deterministic lateraldisplacement at the nanoscale. In this regard, the trajectories offluorescent polystyrene beads with (particle) diameters D_(P)=20-110 nmin a deterministic lateral displacement pillar array were studied. Thenanoparticle experiments allowed not only for an analysis of heretoforeunexplored scales for deterministic lateral displacement, but also tocalibrate and interpret the results of experiments with exosomes (whichhave similar diameters) to be discussed later.

A full description of the design parameters and the nature of theparticle trajectories in the deterministic lateral displacement sortingtechnology has been previously described, for example, in Huang.Briefly, the pillar pitch λ, the row-to-row shift δ, and the gap sizebetween pillars G, (FIG. 3a ) define a critical diameter D_(C). FIGS.3a-f illustrate nanoscale deterministic lateral displacement sortingusing pillar array chips with gap size G ranging from 25-235 nm andmaximum angle θ_(max)=5.7 degrees. At the microscale, particles withdiameter D_(P)>D_(C) will be laterally displaced across an array in“bumping” mode, with a maximum angle θ_(max) (FIG. 3a ) defined by thegeometry of the array as tan(θ_(max))=δλ⁻¹. Particles with diameterD_(P) below D_(C) follow the direction of laminar flow in “zigzag” mode,and thus the overall direction of the trajectory is the same as thelaminar flow, with a mean angle of zero with respect to the array. Inorder to study the properties of nanoscale deterministic lateraldisplacement, arrays were fabricated with gap sizes as small as 25 nm(FIG. 3b ). Additional description of the fabrication details areprovided below.

FIG. 3c shows the capability of the present nanoscale deterministiclateral displacement arrays to displace particles down to 20 nm indiameter using 42 nm gaps. A transitional mode was found between bumping(θ=θ_(max)) and zigzag (θ=0), which is referred to herein as “partialdisplacement mode” where 0<θ<θ_(max) and is represented by thepercentage maximum angle P=θθ_(max) ⁻¹×100. As seen in FIG. 3c , partialdisplacement occurs for 50 nm beads going from G=235 nm to 118 nm. Thisdemonstrates that particles of different particle diameter D_(P) willdisplay different θ for a given G. The efficiency of the displacementprocess can be calculated from η=tan(θ)/tan(θ_(max)) (see FIG.12—described below). For the small migration angles obtained, thisapproximates to η˜θθ_(max) ⁻¹=P/100, so the percentage maximum angleroughly describes the efficiency of sorting in the array. The partialdisplacement mode can be viewed then as a lower efficiency bumping mode.

To explore the sorting capability of deterministic lateral displacementarrays at the nanoscale, a series of monodisperse fluorescence polymerbeads were tested in arrays of gap size G=42 nm, 118 nm, 134 nm, 214 nm,and 235 nm, using the design shown in FIG. 3a . Each particle solutionwas introduced across the entire width of the array inlet and particledisplacement was observed using epifluorescence microscopy. The arraywidth, W=36 μm, and length, L=360 μm, were chosen according to criteriaborrowed from the microscale, such that a particle with diameter aboveD_(C) that enters the array at the top left point of the inlet (seeleftmost panel in FIG. 3d ) would exit the array at the bottom rightpoint of the outlet, following a maximum angle θ_(max)=5.7 degrees. Thismaximum angle was chosen because it combines a relatively large integervalue of row shift repeats (N=10) and high displacement efficiency (seeInglis). It has been shown that D_(C) decreases with N, and thereforethe larger N is, the smaller the diameters of particles that could bedisplaced with maximum angle θ for a given G (see Huang and Inglis).When lateral displacement occurs, the particle density moves towards thecollection wall along the length of the array, generating a fluorescent,triangular pattern with a migration angle, θ (FIG. 3d ). This permitsimaging populations of smaller particles that cannot be trackedindividually (especially for D_(P)<50 nm).

FIG. 3d is an optical microscope image, 20× magnification, of anexemplary nanoscale deterministic lateral displacement device (labeled“nanoDLD array”), showing the overall configuration of the array. FIG.3e provides scanning electron microscope (SEM) images of inlet andoutlet regions bordering the array. FIG. 3f provides fluorescencemicroscopy images of fluorescent polystyrene beads flowing into theinlet region (top row) and exiting the array outlet region (bottom row),corresponding to those shown in the SEM images in FIG. 3e . The lateraldisplacement modes for zigzag, partial, and bumping are shown forD_(P)=20 nm/G=214 nm, D_(P)=50 nm/G=134 nm, and D_(P)=110 nm/G=235 nm,respectively. The migration angle θ, indicates the lateral displacementof the particle flux in the array.

In T. Kulrattanarak et al. “Analysis of mixed motion in deterministicrachets via experiment and particle simulation,” Microfluid Nanofluid10:843-853 (2011) (hereinafter “Kulrattanarak”), the contents of whichare incorporated by reference as if fully set forth herein, displacementangles in between 0 and θ_(max) were observed, and it was concluded thatthis mixed motion between zigzag and bumping was due to the anisotropicpermeability of the arrays. While this effect can still apply tonanoscale deterministic lateral displacement arrays, the low Pecletnumber regime makes the two systems rather different, and diffusion islikely the dominant effect.

Comparison of polystyrene bead displacement with the theoreticalcritical diameter (as determined by Inglis) shows that for particlediameter D_(P)=50-110 nm, the onset of the bumping mode is in agreementwith the existing model. At the row-shift fraction ε=0.10 used in thepresent arrays, Inglis predicts a theoretical critical diameterD_(C)=0.37G, whereas complete displacement was obtained herein at˜0.4-0.47G, which is in agreement with the experimental observations inInglis. The transition across the theoretical critical diameter inaccordance with the present techniques is smoother than the abrupttransition observed for deterministic lateral displacement at themicroscale, consistent with a mixed motion or partial displacement mode(see Kulrattanarak). In contrast to the larger beads, 20 nm beads onlydeflect roughly 32% of the maximum angle. For G=42 nm and D_(P)=20 nm,the Peclet number Pe is Pe˜0.58, assuming an average flow of 300 μm·s⁻¹and a particle diffusivity of D=21.9 μm²·s⁻¹. This small Pe valueimplies that the beads are diffusing across the streamlines, even withina single row shift. This suggests a limit has been approached at whichthe deterministic lateral displacement enters a regime that deviatesfrom the mode of operation at the microscale. Even though at smallPeclet numbers nanoscale deterministic lateral displacement is notstrictly deterministic, the fact that the displacement angle is not zerohas usefulness in potential sorting of nanoscale colloids. In practicalapplications, any partially displacing particle can be fully displacedby increasing the length of the array. A full-displacement length L_(C),can be defined for a given θ and W, at which a partially displacedsample will completely laterally displace (see FIG. 12—described below).This implies that partial displacement can be used to make arrays thatact as “prisms”; splitting a distribution of particles sizes intodifferent angles that can be collected sequentially down the array. Theadvantage of this is that only a single gap size G is needed tofractionate a band of particle sizes.

The dimensions of nanoscale deterministic lateral displacement arraysare compatible with the scales of biological entities ranging fromviruses to small protein aggregates. After demonstrating the possibilityof partial displacement—even at low Peclet numbers, biological colloidswere tested with calibrated fluorescent beads to demonstrate theapplicability of nanoscale deterministic lateral displacement tobiological systems. Many nanoscale bio-colloids exhibit sphericalmorphology and would be expected to show similar displacement behavioras polymer beads, based on the physics nanoscale deterministic lateraldisplacement technology. The present system was tested on monodisperseH1N1 influenza virus and polydisperse human placental mesenchymal stemcell (MSC) derived exosomes. H1N1 influenza A virus has a sphericalvirion of D_(P)=100 nm. See FIG. 4a —which is a schematic diagram of anH1N1 viral particle of 100 nm diameter with fluorescence detectionscheme. Exosomes display a range of particle sizes, D_(P)˜10 s-100 s nm.See FIG. 4d —which is a schematic diagram of a human placental exosomewith fluorescence detection scheme. Here, the migration angle ofdeactivated H1N1 viral particles and human placental mesenchymal stemcell (MSC) derived exosomes were compared to the fluorescent beadcalibration results displayed in FIG. 3c (described above).

H1N1 virus was labeled with a fluorescent antibody and introduced into aG=214 nm array. In the virus free control, the fluorescent antibodies,which are approximately 10 nm in diameter, exhibited a zigzag modebehavior with 0=0 (see FIG. 14b —described below). Displacement ofantibody-virus complexes was observed at θ=θ_(max)=5.7° (see FIG. 4b ),as expected for a D_(P)˜100-120 nm particle. See FIG. 4b —which is acomposite fluorescence microscopy image showing displacement of H1N1viral particles labeled with an Alexa-488-tagged α-N1 antibody in theG=214 nm array. This suggests that viral particles perform similarly topolystyrene beads in the nanoscale deterministic lateral displacementarray. FIG. 4c is a histogram of percentage maximum angle across theoutlet of the array shown in FIG. 4 b.

In contrast, when MSC exosomes are introduced into a G=235 nm array, adispersion of migration angles is observed. See FIGS. 4e and 4f . FIG.4e is a Composite fluorescence microscopy image showing displacement offluorescently labeled exosomes in a G=235 nm array and FIG. 4f is ahistogram of percentage maximum angle across the outlet of the arrayshown in FIG. 4e . Based on the polystyrene bead size calibration, theexosome population can be related to three diameter ranges. Exosomeswere visualized using a lipid-bound fluorescent dye which incorporatesinto the vesicle membrane and does not affect their diameters D_(P).Assuming the exosomes behave like the polystyrene beads, from thecalibration data it was calculated that 1.5% of the exosome populationhas D_(P)>110 nm, 56.3% of the population is between 50 nm<D_(P)<110 nm,and 42.2% of the population has D_(P)<50 nm. See FIG. 4f . Of thislatter population, 69.2% have P>0. Negative P values are due toparticles with trajectories that carry them away from the collectionwall, against the asymmetry of the array. Since these trajectories aredeviations greater than even a zigzag mode, they may be due to a higherdiffusion coefficient for these smaller particles causing randomizedpercolation through the array. The binning of exosome sizes is meant toguide the eye to show the potential for fractioning exosomes; it isexpected that each size exhibits a distribution of migration angles (dueat least in part to diffusion) and therefore the different sizepopulations may overlap to some degree. The comparison implies thatfractions of exosome sizes could be collected by selectively channelingoff particles from the array, opening the potential for probing exosomebiochemistry as a function of particle size. Exosomes with D_(P) up to600 nm would be expected for the samples tested; however, particlesgreater than the gap size, G=235 nm, are filtered out upstream to reduceclogging in the array.

Deterministic lateral displacement arrays have been primarily applied tospherical particles, but there is also interest in sorting polymers,particularly biopolymers such as nucleic acids, by size. Polymers coilinto a compact globular state and it is of interest to understand howthis state behaves in nanoscale deterministic lateral displacementarrays. In that regard, varying lengths of dsDNA, labeled with YOYO-1fluorescent dye, were tested in G=235 nm nanoscale deterministic lateraldisplacement arrays. As in the case of polystyrene beads, a partialdisplacement mode was observed. The migration angle θ, varies as afunction of dsDNA length, appearing to saturate at P=100% around 4 kb.

FIGS. 5a-c are diagrams illustrating DNA displacement in the presentnanoscale deterministic lateral displacement array. Namely, FIG. 5a arefluorescence image mosaics of 0.5-48.5 kb double stranded DNA (dsDNA)displacing in a G=235 nm array. FIG. 5b are normalized fluorescenceintensity line profiles of displaced DNA taken at the outlet (dottedwhite line in FIG. 5a ) of each array. Circles are used in FIG. 5b todenote the inflection point at which the migration angle was measured.FIG. 5c is a diagram illustrating the percentage maximum angle of dsDNAas a function of DNA length.

As the length of the dsDNA molecules are comparable to the dimensions ofthe array, a De Gennes model was used to calculate the confinedend-to-end coil diameter, 2R_(DG) (see FIG. 15—described below). Adescription of the De Gennes model is provided, for example, in M. Daoudet al., “Statistics of Macromolecular Solutions Trapped in Small Pores,”Journal de Physiques, 38 (1), pp. 85-93 (January 1977), the contents ofwhich are incorporated by reference as if fully set forth herein. Forthe calculation, a persistent length of 50 nm was used forYOYO-1-stained dsDNA. Comparing with the expected critical diameter,D_(C)=87 nm, it was seen that 1.0 kb dsDNA has 2R_(DG)=74 nm, so onewould expect to see roughly 50% dsDNA bumping and more than that abovethis chain length, which is what is observed from the displacementexperiments. This would suggest that the displacement condition can bepredicted for the dsDNA, assuming the molecule forms a “particle” ofD_(P)˜2R_(DG). Although the onset of bumping is close to the expectedcritical diameter, only at 4.0 kb, where 2R_(DG)˜295 nm, completedisplacement (i.e., bumping mode) was obtained, thus showing that dsDNAexhibits the same partial displacement behavior as seen in thefluorescent beads. For the two smallest DNA molecules tested, 250 and500 base-pairs (bp), the molecular length is smaller than the gap size,so the de Gennes model does not apply. However, collectively theseresults demonstrate the ability to fractionate dsDNA by size.

In summary, these experiments demonstrate a breakthrough in theapplication of deterministic lateral displacement technology at thenanoscale. Manufacturable silicon processes were used to produce nanoDLDarrays of highly uniform gap sizes, ranging from 25-235 nm. Theseprocesses are compatible with complementary metal oxide semiconductor(CMOS) integration, allowing focus on the next challenge of integratingdigital logic with nanofluidic devices. Using fluorescent nanoparticlesit is demonstrated herein that even at Peclet numbers of order 1, wherediffusion and deterministic displacement compete, nanoDLD arrays can beused to separate particles based on size. Finally, proof of principle ofsize-based separation of exosomes, viruses and DNA has been shownherein. The nanoDLD array constitutes a building block in a newgeneration of on-chip fluidic processing technologies that couldpotentially be multiplexed to produce improved on-chip diagnostics.

A description of the array and particle preparation, as well as furtherdetails on the analysis of the nanoscale deterministic lateraldisplacement fluorescence images is now provided

Fluidic Chip and Pillar Array Fabrication:

Nanofluidic chips were fabricated on 200 mm wafers to enablehigh-quality fluorescence imaging of nanoscale polymer beads andbio-colloids when coupled with a custom fluidic jig as described, forexample, in Wang et al., “Hydrodynamics of Diamond-Shaped GradientNanopillar Arrays for Effective DNA translocation into Nanochannels,”ACS NANO, vol. 9, no. 2, pp. 1206-1218 (January 2015) (hereinafter“Wang”), the contents of which are incorporated by reference as if fullyset forth herein. See FIG. 6a —which is an image of these fluidic chipsprinted on a 200 nm wafer using mixed mid-ultraviolet (MUV) and electronbeam (e-beam) lithography. Optical contact lithography followed by acombination of e-beam and deep-ultra violet (DUV) lithography schemes(see, for example, Wang et al., “200 nm Wafer-Scale Integration ofSub-20 nm Sacrificial Nanofluidic Channels for Manipulating and ImagingSingle DNA Molecules,” 2013 IEEE International Electron Devices Meeting(IEDM) (December 2013), the contents of which are incorporated byreference as if fully set forth herein) were utilized consecutively todefine microchannel and nanofluidic pillar array features, respectively,in an silicon dioxide (SiO₂) hard mask on bulk silicon substrates. SeeFIG. 6b . Following hard mask definition, all features were transferredinto silicon in tandem using a reactive-ion etch (RIE).

40×40 mm square fluidic chips (12 per wafer) were fabricated on 200 mmSi wafers with a blanket layer of thermal oxide ranging from 100-150 nm.Oxide thickness was selected depending on the target depth of thesilicon etch corresponding to a particular pillar height. An opticalcontact lithography SUSS MA8 mask aligner was used to print microchannelfeatures in a photoresist mask, which was subsequently etched into SiO₂with an etch stop on the Si bulk using RIE (See FIG. 6a ).

After defining microchannels in the SiO₂ hard mask, a negative-tone,e-beam lithography strategy was employed to provide the sharpestpossible resolution of the nanofluidic pillar array features, connectingsets of microchannels (see FIG. 6b ), and a positive DUV-definedlithography window was used to mask the remainder of the wafer outsidethe nanofluidic pillar array region from RIE processing when definingthese features in the thermal oxide hardmask. E-beam lithography wasperformed on a high-resolution VectorBeam 6 (Leica VB6HR) system todefine pillar arrays in a resist stack consisting of an HM8006 organicplanarization layer (OPL) (JSR) coated with a thin 2% hydrogensilsesquioxane (HSQ) (Dow Corning) negative tone e-beam resist. Afterdeveloping the exposed features, the wafers were coated with a 0.45μm-thick positive photoresist and DUV window regions were printeddirectly on top of the e-beam resist features using a 0.65 NA DUVstepper (ASML) (see FIG. 6b ). RIE was then used to transfer the e-beamfeatures into the thermal oxide hard mask and stop on Si after which allresist was stripped in an oxygen (O₂) plasma asher (GaSonics).

With all features defined in the SiO₂ hard mask, RIE was used tosimultaneously etch the pillar array and microchannel features intosilicon. Optimized RIE processing was carried out in an DPSII ICP etchchamber (Applied Materials, CA) for pattern transfer to fabricate200-450 nm high Si pillars from the e-beam resist pattern, depending onthe desired gap width. First, the negative-tone HSQ resist was used toetch through the OPL using an N₂/O₂/Ar/C₂H₄ chemistry at 400 watts (W)source power, 100 W bias power, and 4 millitorr (mTorr) pressure at 65°C. The SiO₂ hard mask was then patterned using CF₄/CHF₃ chemistry at 500W source power, 100 W bias power, and 30 mTorr pressure at 65° C.

Next, the OPL carbon resist was stripped using O₂/N₂ chemistry in anAxiom downstream asher (Applied Materials, CA) at 250° C. Finally, usingthe SiO₂ hard mask, Si features defining the arrays and microchannelswere etched using the DPS II ICP etch system (Applied Materials, CA) byfirst a CF₄/C₂H₄ breakthrough step and then Cl₂/HBr/CF₄/He/O₂/C₂H₄ mainetch at 650 W source power, 85 W bias power, and 4 mTorr pressure at 65°C. Final chip preparation required stripping the hard mask oxide andregrowing a thin thermal oxide layer ranging from 10-50 nm.

Nanoscale Deterministic Lateral Displacement Chip Preparation:

To produce a functioning nanoscale deterministic lateral displacement(nanoDLD) device from the fabricated array chips, a glass coverslip wasbonded over the array to provide a water-tight enclosure. Chips werebonded as reported in Wang, using the same coverslip design. Bondedchips were annealed at 550° C. for 7 hours in a convection oven(Lindberg/Blue), and stored in a nitrogen dry box until needed. In somecases, bonded chips were further surface functionalized prior toexperiments; (see below). Bonded chips were used in experiments within 2weeks of annealing.

Deterministic Lateral Displacement Chips Wetting Protocol:

To run a deterministic lateral displacement experiment on a bonded chip,the arrays needed to be wetted with fluid. Chips were set vertically ina custom glass holder, with inlets below outlets. The chips weresubmerged in enough ethanol (200 proof, Pharmco-AAPER) so that only theinlet ports were covered, while the outlet ports remained open to air toallow proper wetting. The ethanol was allowed to capillary wet eachfluidic array entirely (˜10-30 min). The chips were then transferred todiH2O (Millipore) and fully submerged for 60 minutes. At this stagechips can be kept in water until needed for experiments without evidenceof de-wetting.

Surface Modification of Nanoscale Deterministic Lateral DisplacementDevices:

For running bio-colloids (i.e., viral particles, exosomes, and dsDNA) itwas determined that surface modification of the array silica was neededin order to prevent sample adhesion to the device surfaces.Bio-colloids, especially exosomes, when run in unmodified chips wouldclog the micro-array inlet and would not reach the array.

For surface modification, a chip was immediately transferred from theoven, after annealing, to a 500 milliliter (mL) glass, cylindrical, flatflange reactor (Wilmad-LabGlass). The chip was positioned upside down(inlets down) on a custom glass holder such that a stir bar couldoperate beneath the chip. The reactor was sealed with a 3-neck head, andpurged with nitrogen flow for 30 minutes Via cannula, 250 mL of a 10millimolar (mM) solution of2-(methoxypoly(ethyleneoxy)6-9propyl)dimethylchlorosilane (technicalgrade, 90%, Gelest Inc.) dissolved in degassed, anhydrous chloroform(Sigma Aldrich), was added to the reactor. The liquid level submergedonly the inlet holes of the chip, allowing capillary action to wicksolvent up into the arrays. Care was taken not to splash and wet theoutlet ports of the chip while loading the reagent, as this leads tobubbles, and unmodified regions in the device. The chip was allowed tosit for 16 hours at room temperature, under nitrogen, with gentlestirring. The chip was then removed and transferred into 300 mL ethanol(200 proof, Pharmco-AAPER) and stirred for 16 hours. This process wasrepeated for diH₂O (Millipore). At this stage, chips could be kept inwater until needed for experiments. Chips have been kept up to 1 monthsubmerged before running successfully.

Particle Sample Preparation:

Polystyrene Beads:

Aqueous, fluorescent polystyrene beads, with carboxylic functionalgroups were purchased from commercial suppliers, with particle diametersD_(P)=20 nm (Molecular Probes, Life Technologies, Thermo FisherScientific, Inc.), 50 nm and 110 nm (Bangs Laboratories, Inc.). Allbeads have absorption bands compatible with 488 nm excitation, andemission at 510-520 nm. Bead samples to be run in arrays were preparedby diluting the as supplied bead solutions in TE buffer (BioUltra, formolecular biology, pH 7.4, Sigma Aldrich) with 3% v/vtotal2-mercaptoethanol (Sigma Aldrich), to inhibit photo-oxidation, and 2-10%v/vtotal Tween20 (For molecular biology, Sigma Aldrich), to preventparticle aggregation and clogging. Typically a sample solution of 200 μLwas prepared.

FIGS. 7a-e show the physical properties of the beads used inexperiments. Namely, FIGS. 7a-c are SEM images of D_(P)=110 nm, 50 nm,and 20 nm, respectively, beads coated with a layer of evaporated Ti/Au(1 nm/10 nm). The scale bars shown represent 200 nm. FIG. 7d is ahistogram of bead diameters measured from the SEM images in FIGS. 7a-c .FIG. 7e is a table of properties of the bead samples used in thenanoscale deterministic lateral displacement experiments. a: Meandiameter±standard deviation.

Bead solutions were momentarily vortexed to mix, and then centrifuged at3000 revolutions per minute (RPM) for about 30 seconds. Bead solutionswere prepared fresh daily for each experiment. Prior to use, 5microliters (μL) of solution were set between two glass microslides andimaged to visually verify the quality of the beads solution.

Exosomes:

Placental mesenchymal stem cell (MSC) derived exosomes were obtainedfrom Zen-Bio, Inc. Mean particle diameter is <DP>=290 nm, 100 μg, 165μL, 0.60 mg·mL-1, 9.5·108 particles·mL-1. Particles were purchasedlabeled with DiO lipophilic cyanine dye (484 nm excitation, 501 nmemission). The as-obtained material was split into 10×, ˜15 μL aliquotsand frozen at −80° C. until needed. Prior to running an experiment, analiquot was removed from cold storage and allowed to thaw at 4° C. for30 minutes. The sample was then vortexed momentarily and thencentrifuged at 3000 RPM for about 30 seconds to recollect. The samplewas directly loaded into the flow chamber on the fluid jig.

Viral particle preparation: Monoclonal antibodies against H1N1neuraminindase (Sigma, SAB3500064) and M13 [E1] (Abcam, ab24229) werelabeled with AlexaR-488 on primary amines (Life Technologies). 33nanomolar (nM) inactivated influenza viral particles (AFLURIA, bioCSL,2014-2015) were incubated with 2.2 nM fluorescent antibody for 30minutes at 4° C. in a solution of 14 mM NaCl, 0.5 mM Na2HPO4, 20 nMKH2PO4, 160 nM KCl, and 2 mM CaCl2. 15 μL of sample was introduced to a212 nm gap array.

DNA Sample Preparation:

Stock solutions of 0.5-20 kb DNA and λ-DNA (0.5 μg/μL, New EnglandBiolabs) were diluted to 100 μg/μL in 10 mM TE buffer (10 mM Tris, 1 mMEDTA, pH 8, Life Technologies) with 3% 2-mercaptoethanol and 0.1% TWEEN20. DNA was labeled with YOYO-1 iodide fluorescent dye (491/509 nm)(Life Technologies) at a DNA base pair-to-dye ratio of 5:1. Samples wereincubated at room temperature for 2 hours, and stored at 20° C. for use.

Running Nanoscale Deterministic Lateral Displacement ParticleDisplacement Experiments:

Bonded and wetted chips were used for running displacement experiments.A chip was loaded into a custom built fluid jig. See FIGS. 8a-b . FIG.8a is a schematic diagram of the fluid jig, and FIG. 8b is a top-downimage of the fluid jig with a nanoscale deterministic lateraldisplacement chip loaded. In the example depicted in FIG. 8b , twelvethreaded ports (six inlets and six outlets) are shown. The rectangularwindow, through which the microscope objective can reach the chip toimage, can be seen in the middle of the jig.

As shown in FIG. 8a , the fluid jig is composed of a mounting base and aconnector plate into/through which high-pressure liquid chromatography(HPLC) fittings can be screwed to inject fluid samples. The connectorplate has a rectangular window designed to allow up to a 100×oil-immersion objective (Zeiss N-Achroplan 100×/1.25 oil, Zeiss,Germany) to be used for fluorescence imaging. The sample is loaded intothe inlet reservoir of the connector plate, and the plate then screweddown onto to the chip/mounting base. A syringe pump (QMixx, Cetoni GmBH,Germany) is connected to the inlet port of the connector plate. The 10mL syringe+tubing is filled with diH₂O (Millipore). Imaging is carriedout on a Zeiss Scope.A1 upright fluorescence microscope coupled with anAndor iXon Ultra 897 (Andor Technology Ltd., Oxford Instruments, UK)EMCCD camera connected to a computer, where both imaging and pumppressure are controlled. A 470 nm light emitting diode LED was used forexcitation, with the Zeiss filter set 38 HE (470/40 excitation, 495 beamsplitter, 525/50 emission).

Array Structures:

Fast Fourier transform (FFT) analysis of the arrays (see FIG. 6c )confirmed a maximum angle θmax, of 5.8° which is very close to thedesign value of 5.7°. The maximum angle θmax=5.7° was chosen as itcorresponds to a relatively small row shift fraction c=0.1 leading to awell-defined row repeat N=10, where θmax=tan⁻¹ (ε) and N=1/ε². Having aprecise N simplifies design criteria while a small c increases theefficiency of a particular gap size in sorting the smallest possibleparticle, since the critical diameter Dc to sort a particle in theparabolic flow model is given by Dc=2αGε with α being a scaling factordepending on the flow profile. This sorting efficiency becomes crucialto induce sorting of very small entities such as proteins, particularlyas gaps are scaled in the tens of nanometers regime, pushing the limitsof fabrication and ability to effectively wet these features.

FIG. 6d shows the cross-sectional SEM image of a G=134 nm pillar arraywith pillar pitch λ=400 nm. Gap scaling and uniformity is demonstratedthrough RIE optimization and thermal oxidation of the Si posts,permitting well-controlled gap widths. Gap sizes for each array testedwere determined by randomly measuring 3 sets of 5 adjacent gaps acrossthe ˜35 μm width of the pillar arrays (a statistical average of 15 gapsper array), including 1 set chosen near the array center and the other 2sets nearer to each edge of the array. Gap variation from pillar top tobottom was minor but not negligible so measurements were taken at pillarmid-height as indicated by the dotted line in FIG. 6d . Thermaloxidation after pillar etching on parallel-processed wafers permittedcontrollable means of tuning of the gap size to effectively narrow thegap to a desired width based on the results from an SEM cross-section ofa send-ahead wafer for each array fabricated.

General Experimental Layout:

When running a sample, a syringe pump is used to control the pressure toobtain a stream flow at 200-300 μm s-1. In typical experiments itrequires 10-30 min for particles to reach the array from the connectorplate inlet. Video images are collected of particle flux across theinlet of the array to assess the degree of particle distribution priorto injection into the array, and across the outlet to capture the degreeof displacement at the end of the process. Exposure time is 17.9 ms, 200frames per video. For each combination of gap size G and particlediameter DP, three or more independent experiments (individual arrays)were run.

In the case of exosomes, a different fluid jig was used (see Wang) whichuses negative pressure through vacuum to drive the fluid flow. Anexosome sample was directly loaded into the inlet fluidic reservoir ofthe flow chamber and a ˜1 torr vacuum applied to the pump connector onthe outlet side of the jig.

Analysis of Nanoscale Deterministic Lateral Diffusion FluorescenceMicroscopy Images for Particle Behavior:

Analysis of Fluorescent Polystyrene Bead Displacement:

Fluorescence image videos of the array outlet are analyzed using customsoftware to determine the migration angle of the displaced particleflux. In the current array configuration, the flux of particles acrossthe array is displaced towards the collection wall, forming afluorescent, triangular pattern (white triangle in FIG. 9a ). Adepletion region, where particles have been displaced out, appears onthe opposite side of the array from the collection wall (dark trianglein FIG. 9a ). The extent of this depletion region at the outlet of thearray determines the lateral displacement ΔW, of the particle flux.Determination of the lateral displacement is complicated by the factthat the edge of the particle flux, seen in the cross-section offluorescence intensity across the array outlet, has a continuous formwith no sharp cut-off. The “edge” of the particle flux is estimated, andthus ΔW, using the inflection point of the fluorescence intensity (FIG.9b ). This assumes that the fluorescent intensity distributioncorresponds to the particle density distribution. As shown in FIG. 9a ,particles are displaced upwards towards the collection wall of thearray, forming a fluorescent triangle pattern (wedge), from which themigration angle θ, and lateral displace, ΔW, can be measured. Thelateral displacement is taken at the outlet of the array. In theschematic, the particles are completely displacing (bumping mode) soθ=θ_(max)=5.7°, the maximum angle of the arrays used in this work. InFIGS. 9b and 9c , the dashed lines show the 1st derivative of thefluorescent line profiles, indicating the inflection point (black dot).The lateral displacement, ΔW, is taken as the distance between theleft-most wall of the array (opposite the collection wall) and theinflection point. Using the length of the array, L, and the lateraldisplacement, ΔW, the migration angle can be calculated fromtan(θ)=ΔW/L. The line which is the 1st derivative taken before a50-point smooth of the data (dashed line) is labeled in FIG. 9 c.

The migration angle θ, is defined as tan(θ)=ΔW L⁻¹, where L is thelength of the array. For a completely displaced particle sample, allparticles end up at the collection wall at the end of the array, andθ=θ_(max)=5.7°, the maximum angle of the array. For no displacement, theparticle flux covers the entire outlet, and θ=0. The displacementefficiency is defined as:

$\eta = {\frac{\Delta \; W}{W} = {{\frac{\tan \; \theta}{\tan \; \theta_{\max}} \sim \frac{\theta}{\theta_{\max}}} = \frac{P}{100}}}$

To compare the effectiveness of sorting particles for a given DP and G,a figure of merit, FOM, is defined as the ratio between the lateraldisplacement of the particles, and the distance needed to fully displacethe particles across the array:

FOM=tan θ=η tan θ_(max)

From the definition of the figure of merit, the displacement length canbe defined as:

$L_{C} = \frac{W}{F\; O\; M}$

FIG. 10 is a diagram illustrating polystyrene fluorescent beaddisplacement as a function of particle diameter, D_(P), compared tocritical diameter needed for displacement in a parabolic flow. Beadvalues are given for a given row shift fraction, ε=0.10, and scalingratio D_(O)G⁻¹. Value shading represents the percentage maximum angle,P. The black line is the calculated critical diameter scaling ratio,D_(C)G⁻¹=1.16ε^(0.5). Theoretically, beads with scaling ratios below thecritical line should exhibit zigzag mode, P=0%, and not displace withinthe array, while those above should show complete displacement, P=100%.

FIG. 11 is a diagram illustrating percentage maximum angle offluorescent polystyrene beads displaced in nanoscale deterministiclateral displacement arrays as a function of the scaling ratio betweenparticle diameter and gap size. Bead diameters are 110 nm (squares), 50nm (circles) and 20 nm (triangles). Error bars represent the standarddeviation of at least three independent experiments. The line atD_(P)G⁻¹=0.37 represents the theoretical critical diameter, D_(C), inparabolic fluid flow at which beads are expected to be in bumping mode.P=100% corresponds to complete displacement of beads (bumping mode),P=0% corresponds to no displacement (zigzag mode), and 0%<P<100%indicates partial displacement mode.

FIG. 12 is a table of performance parameters for nanoscale deterministiclateral displacement of fluorescence polystyrene beads.

Analysis of Exosome and Virus Displacement:

For exosomes and viral particles, single-particle trajectories arerecorded in fluorescence microscope images, instead of a flux density,as in the case of fluorescent polystyrene beads. This means that adistribution of single-particle events were obtained, instead of acontinuous distribution determined by the average fluorescence density.In general, flowing particles form a streak or “trace” in a given videoframe. For each particle observed, the location of the head of theparticle's trace is manually marked per frame of video. The collectionof x,y-coordinate pairs taken from the combined number of frames(typically 200) defines the trajectory of the particle within the imageframe of the video (see FIGS. 13a-c ).

The migration angle θ, is defined as tan(θ)=ΔX/ΔY, withΔX=_(xfinal-xinitial) and ΔY=_(yfinal-yinitial), using the initial andfinal x,y-coordinates of the particle trajectory. In determining θ, onlytrajectories that initiate at a distance larger than 10% of the arraywidth from the collection wall are used, in order to avoid thepossibility of miscounting particles that are partially displacing asones which are bumping. The value of 10% of W comes from analysis of thehalf-width at half-maximum of the 110 nm bead fluorescence intensityagainst the collection wall, which shows complete bumping mode. From thecollected θ of all the single-particle trajectories, a histogram of thepercentage maximum angle P, can be generated. This distribution ofangles is equivalent to the fluorescence intensity line profiles used inthe bead analysis, however it is acquired from the accumulation ofsingle-particle data rather than the measurement of an ensemblefluorescence.

FIG. 13a is a fluorescent microscopy image of an exosome particle. Aseries of 6 images, taken every 36 milliseconds, overlaid together showsthe trajectory of the particle through the nanoscale deterministiclateral displacement array. The particle appears as a small line (trace)due to the finite exposure time (18 milliseconds). Scale bar represents10 μm. The single-particle trajectory can be measured from the movementof the head of the trace through the array as a function of time. FIG.13b illustrates collection of single-particle exosome trajectories takenat the outlet of a G=235 nm array. Traces in dotted white line initiatedwithin 10% of the array width from the collection wall and are not usedfor determining migration angle, while those in black initiate outsidethis threshold. FIG. 13c is a histogram of single-particle positions atthe outlet (black horizontal line in FIG. 13b ). Values in dotted whiteline and black correspond to those in FIG. 13b . The migration angle canbe calculated from the amount of lateral displacement (x-position) ofthe particle from the start of the trajectory, and the length ofdistance travelled (y-position, FIG. 13b ).

Control samples of H1N1 virus alone and fluorescently labeled anti-H1N1antibody alone were run through the array. See FIGS. 14a-c .Additionally, a non-H1N1 virus binding fluorescently labeled anti-M13antibody with virus was run through the array. This demonstrates thatbumping mode is only observed when both virus and a specific antibodyare present.

Specifically, FIGS. 14a-d are images of viral particle experimentalcontrols—i.e., representative composite fluorescent microscope image ofAlexa Fluor® 488-labeled anti-H1N1 neuraminidase and M13 antibodies andinactivated influenza viral particles in a 212 nm gap arrayfunctionalized with a polyethylene glycol silane ligand. Direction offlow is from top to bottom with particles migrating towards thecollection wall on the right of the image. The image is composed ofsequential frames showing antibody-virus complex trajectory (frametime=18 ms). Virus alone shows no fluorescence (see FIG. 14a ).Antibodies alone follow the streamlines of the laminar flow (see FIG.14b ) while anti-H1N1 antibody-virus complexes are bumped (see FIG. 14c). Non-specific M13 antibodies do not bind viral particles and thereforedo not bump (see FIG. 14d ).

Analysis of dsDNA Displacement:

dsDNA experiments were analyzed using the same methods as fluorescentpolystyrene beads (see above). The de Gennes mean confined coil radiuswas calculated using:

$R_{DG} = {{Nh}\frac{({wp})^{1/3}}{D^{2/3}}}$

With persistence length, p=50 nm, molecular width, w=2.4 nm, length perbase, h=0.34 nm, number of bases per strand, N, and geometric average ofnanochannel taken as D=(GH)^(1/2)=307 nm, with nanopillar gap size G=235nm and gap height H=400 nm.

FIG. 15 is a table of end-to-end coil diameters and scaling ratioscalculated for dsDNA in nanoscale deterministic lateral displacementexperiments.

Although illustrative embodiments of the present invention have beendescribed herein, it is to be understood that the invention is notlimited to those precise embodiments, and that various other changes andmodifications may be made by one skilled in the art without departingfrom the scope of the invention.

What is claimed is:
 1. A method for virus detection, comprising thesteps of: collecting a fluid sample from a virus-bearing source;contacting the fluid sample with an antibody that binds to viruses toform a sample-antibody mixture, wherein the antibody is labeled with afluorescent tag; separating particles including any antibody-viruscomplexes, if present, from the sample-antibody mixture using an assaynanopillar array; and detecting the antibody-virus complexes, ifpresent, in the particles from the separating step using fluorescence.2. The method of claim 1, wherein the virus-bearing source is selectedfrom the group consisting of: blood, saliva, sweat, plant tissue,drinking water, and food products.
 3. The method of claim 1, wherein thefluorescent tag is selected from the group consisting of: quantum dots,Alexa Fluors®, fluorescein, rhodamine, Oregon green, pyrene, and HyLite™Fluor dyes.
 4. The method of claim 1, wherein the assay nanopillar arraycomprises a 120 nanometer nanopillar array
 5. The method of claim 1,further comprising the step of: removing particles from thesample-antibody mixture using a filtering nanopillar array prior to theseparating steps.
 6. The method of claim 5, wherein the filteringnanopillar array comprises a 1 micrometer nanopillar array.
 7. Themethod of claim 1, wherein the step of contacting the fluid sample withthe antibody comprises the step of: mixing the fluid sample with asolution containing the antibody labeled with the fluorescent tag. 8.The method of claim 1, further comprising the step of: placing theantibody labeled with the fluorescent tag in a mixing reservoir.
 9. Themethod of claim 8, wherein the step of contacting the fluid sample withthe antibody comprises the step of: passing the fluid sample through themixing reservoir whereby the fluid sample contacts the antibody.
 10. Themethod of claim 1, wherein the antibody-virus complexes, if present, inthe particles are detected using a diode-induced fluorescence detector.11. The method of claim 1, wherein an amount of the fluid samplecollected is from about 50 microliters to about 100 microliters, andranges therebetween
 12. A virus detection chip device, comprising: acapillary opening for accepting a fluid sample collected from avirus-bearing source; a mixing reservoir, connected to the capillaryopening, for contacting the fluid sample with an antibody that binds toviruses, wherein the antibody is labeled with a fluorescent tag to forma sample-antibody mixture; a first nanopillar array, connected to themixing reservoir, for removing particles from the sample-antibodymixture; a second nanopillar array, connected to the first nanopillararray, for separating particles including any antibody-virus complexes,if present, from the sample-antibody mixture; and a diode-inducedfluorescence detector, connected to the second nanopillar array, fordetecting the antibody-virus complexes, if present, in the particlesusing fluorescence.
 13. The virus detection chip device of claim 12,wherein the virus-bearing source is selected from the group consistingof: blood, saliva, sweat, plant tissue, drinking water, and foodproducts.
 14. The virus detection chip device of claim 12, wherein thefluorescent tag is selected from the group consisting of: quantum dots,Alexa Fluors®, fluorescein, rhodamine, Oregon green, pyrene, and HyLite™Fluor dyes.
 15. The virus detection chip device of claim 12, wherein thefirst nanopillar array comprises a 1 micrometer nanopillar array. 16.The virus detection chip device of claim 12, wherein the secondnanopillar array comprises a 120 nanometer nanopillar array.
 17. Achip-based immunoassay method, comprising the steps of: providing avirus detection chip device having: a capillary opening; a mixingreservoir connected to the capillary opening; a first nanopillar arrayconnected to the mixing reservoir; a second nanopillar array connectedto the first nanopillar array; and a diode-induced fluorescence detectorconnected to the second nanopillar array; introducing a fluid samplecollected from a virus-bearing source to the virus detection chip devicethrough the capillary opening; in the mixing reservoir, contacting thefluid sample with an antibody that binds to viruses, wherein theantibody is labeled with a fluorescent tag to form a sample-antibodymixture; removing particles from the sample-antibody mixture using thefirst nanopillar array; separating particles including anyantibody-virus complexes, if present, from the sample-antibody mixtureusing the second nanopillar array; and detecting the antibody-viruscomplexes, if present, in the particles using the diode-inducedfluorescence detector.
 18. The method of claim 17, wherein thevirus-bearing source is selected from the group consisting of: blood,saliva, sweat, plant tissue, drinking water, and food products.
 19. Themethod of claim 17, wherein the first nanopillar array comprises a 1micrometer nanopillar array.
 20. The method of claim 17, wherein thesecond nanopillar array comprises a 120 nanometer nanopillar array.